INTRODUCTION

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Corynebacterium glutamicum is an important industrial amino-acid-producing bacterium. This organism is known to possess very high intracellular concentrations of glutamic acid, i.e., up to 200 mM (24). In microorganisms, glutamate can be synthesized by two alternative pathways (12). NADP-dependent glutamate dehydrogenase (GDH) catalyzes the reductive amination of 2-oxoglutarate to glutamate (GDH pathway). Glutamate can also be formed by the consecutive reactions of glutamine synthetase (GS) and glutamine 2-oxoglutarate-amidotransferase (GOGAT) (i.e., the GS/GOGAT pathway). GS catalyzes the ATP-dependent amination of glutamate to glutamine. GOGAT catalyzes the reductive transfer of the -amidogroup of glutamine to 2-oxoglutarate with concomitant oxidation of NADPH. All of these ammonium-assimilating enzymes have been described in amino-acid-producing bacteria (41, 46, 50), indicating that both pathways of ammonium assimilation may function simultaneously in C. glutamicum.

It has long been assumed that C. glutamicum requires GDH as a key enzyme for NH₄⁺ assimilation during glutamate production. However, a GDH mutant of C. glutamicum was recently constructed, and it was shown that neither growth nor glutamate production was impaired in this mutant (4). While it seems plausible to assume that the GS/GOGAT pathway was responsible for ammonium assimilation in that case, no detailed tests to verify whether alternative (e.g., alanine dehydrogenase) routes may be active were performed.

While determination of enzyme activities in crude extracts may give a first clue as to which ammonium assimilation pathways are active, they cannot be used to predict the in vivo flux distribution over competing enzyme systems such as GDH and GS/GOGAT.

In vivo nuclear magnetic resonance (NMR) spectroscopy, especially when used in combination with stable isotope labeling, does allow the characterization of metabolic activities in the living cell (34, 42). With ¹⁵N-NMR, important metabolites such as glutamate, glutamine, alanine, lysine, aspartate, and N-acetylglutamate have been detected in bacteria, fungi, and algae (13, 25, 27). The relative importance of especially the GDH and the GS/GOGAT pathways in NH₄⁺ assimilation has been studied successfully by using the ¹⁵N-NMR technique in, e.g., Bacillis macerans and B. polymyxa (16, 17), B. azotofixans (18), Aspergillus nidulans (26), Clostridium kluyverii and C. butyricum (19), and Agaricus bisporus (2). All of these studies used cell extracts taken at different time points after incubation with ¹⁵N-labeled substrates to identify the principal metabolites and pathways involved in ¹⁵N assimilation in a mainly qualitative manner. In contrast, detailed quantitative studies of nitrogen flux distribution in vivo in microorganisms have not yet been reported.

Although the analysis of carbon fluxes in the central metabolism by metabolite balancing (44, 51) and computer-aided stationary-state isotopic analysis of ¹³C-labeled intermediates (30, 54, 55) nowadays may be considered as an established technique that allows the differentiation between parallel pathways (31, 43) and even the identification of bidirectional metabolic fluxes (30, 48, 54, 55), it is not well suited for nitrogen flux analysis. This is due to the fact that nitrogen labeling of metabolic intermediates upon prolonged incubation with a one-nitrogen substrate (e.g., urea, ammonium, glutamate, or aspartate) does not show any positional effects, i.e., at isotopic steady state the nitrogen enrichment in all intermediates is equal to that of the substrate, independent of the fluxes. Although the use of extensive modelling of glutamate metabolism (29) allows determination of some nitrogen fluxes (e.g., GDH [8]) on the basis of glutamate ¹³C enrichments, a complete analysis of the ammonium assimilation flux network as envisaged for the present study requires time-resolved measurements of ¹⁵N label incorporation kinetics.

While such experiments were hitherto practically impossible due to the very low sensitivity of conventional ¹⁵N-NMR techniques, which do not allow for a useful time resolution in dynamic metabolic studies, recent developments in the field of integrated NMR and fermentation equipment fortunately have opened new perspectives. A continuous-flow NMR bioreactor enabled monitoring of the metabolism of anaerobic Zymomonas mobilis with in vivo ³¹P-NMR at fourfold-increased sensitivity compared to conventional NMR approaches (9). A further development of this system resulted in a hydrocyclone bioreactor that permitted the continuous aerobic cultivation of C. glutamicum at a cell density of 25 g (dry weight)/liter allowing the metabolism to be monitored with in vivo ¹³C-NMR at a time resolution of 10 min (14). In view of the high glutamate pool in C. glutamicum, this system seemed also to offer good perspectives for application of in vivo ¹⁵N-NMR.

This study is concerned with the detailed quantitative investigation of ammonium assimilation in C. glutamicum wild type (ATCC 13032) and its GDH mutant. The aim was to determine over which pathways these C. glutamicum strains assimilate ammonium and to quantify the primary nitrogen fluxes in both strains in vivo.

	MATERIALS AND METHODS

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Bacterial strains and growth conditions. The experiments were performed with C. glutamicum wild type (ATCC 13032) and a GDH-negative mutant C. glutamicum EB1 (4). Continuous cultures were run with a fully synthetic medium as described elsewhere (20), modified for the glucose concentration (100 g/liter) and the (NH₄)₂SO₄ concentration, which was varied between 10 and 15 g/liter by separate dosage. The medium was inoculated with washed cells from an overnight culture in BHI medium (Difco) to give an initial optical density at 600 nm (OD₆₀₀) of about 5. Fermenter operation in the continuous cultivation mode was started directly after inoculation. Thus, growth to high cell densities took place during continuous culture operation and not in a batch phase. All of the continuous culture fermentations reported here were performed in vivo in the NMR bioreactor at a high cell density of 50 g (dry weight)/liter. The medium dilution rate was set at 0.1 h¹, and the growth rate was set to 0.05 h¹ by adjusting the bleed flow. In the fermentations, pH was controlled at 7.0 by using 2 M NaOH, and the temperature was kept at 30°C. The pO₂ was regulated at 20% of air saturation. The continuous cultures were aerated with a 50% (vol/vol) N₂-O₂ mixture to reduce gas input to the fermentation broth.

Preparation of crude extracts and enzyme assays. Cells from 2 ml of culture broth were harvested by centrifugation (10 min, 6,000 × g), subsequently washed two times in 40 ml of washing buffer solution (for specifications, see below), and resuspended in 1 ml of sonication buffer (for specifications, see below). Sonication (UP 200 S sonifier; Dr. Hielscher GmbH, Teltow, Germany) was performed at 0°C for 5 min. Cell debris were removed by centrifugation (13,000 rpm, 30 min, 4°C), and the resulting cell extract was used for enzyme assays. The protein concentration was determined by the biuret method with bovine serum albumin (Boehringer Mannheim) as the standard.

NMR spectroscopy. All NMR experiments were performed by using an AMX-400 WB spectrometer system (Bruker, Karlsruhe, Germany).

(i) In vivo ¹⁵N-NMR. Cells were cultivated in a continuous-flow membrane bioreactor system equipped with a hydrocyclone reactor vessel adapted for defined bypass volume flow as described before (14). NMR experiments were started after at least 50 h of continuous cultivation. The reactor vessel was introduced into the magnet approximately 1 h before the start of the ¹⁵N experiments. To reduce the amount and size of the gas bubbles in the NMR-sensitive region, the system flow was then increased to 800 liters/h, resulting in a better gas-liquid separation. ¹⁵N-NMR was carried out at 40.5 MHz by using a custom-tailored 20-mm ³¹P-¹³C-¹⁵N-NMR probehead (Bruker Spectrospin, Faellanden, Switzerland). Shimming was performed under flow conditions by using the water signal measured with the detuned ¹H decoupling coil. Line widths of 10 to 12 Hz for ¹⁵N were routinely achieved. The extremely long ¹⁵N longitudinal relaxation times (the T₁ of ammonium-¹⁵N, for example, was determined to be ca. 50 s) forced use of continuous broadband ¹H decoupling, exploiting the large (but negative) ¹⁵N nuclear Overhauser effect (NOE). Although under our continuous-flow conditions only a fraction of the theoretically maximal NOE can build up inside the NMR measurement chamber in the sensitive volume of the decoupling coil, sensitivity was still much better than without the NOE. The following ¹⁵N pulsing conditions were found to be optimal: 90° pulses (70 µs) with a recycle delay of 200 ms, 20.8-kHz sweep width, 2,048 complex data points, continuous Waltz-16 composite pulse broadband proton decoupling (90° ¹H pulse, 200 µs). A total of 4,440 and 2,220 scans per spectrum were used for wild-type and GDH mutant strains, resulting in a time resolution of 15 and 7.5 min, respectively. One initial ¹⁵N spectrum was recorded just before the addition of the labeled ammonium. The recording of a series of consecutive ¹⁵N spectra (a 3-h total NMR measurement time) was started immediately after application of a single pulse of (¹⁵NH₄)₂SO₄ (>99% ¹⁵N; Cambridge Isotope Laboratories). The spectra were processed without line broadening. Peak areas were determined by using the deconvolution package of the UXNMR spectrometer software (Bruker, Karlsruhe, Germany).

(ii) ¹⁵N-NMR of ethanolic cell extracts for calibration. Samples of cell suspension (10 ml) were drawn from the reactor for amino acid pool size quantification and ¹⁵N-NMR analysis at the following time points after addition of the labeled (¹⁵NH₄)₂SO₄: 1, 2, 3, 4, 5, 10, 15, 20, 25, 30, 45, 60, 75, 90, 105, 120, 150, and 165 min. This loss of biomass from the system was partially compensated for by the cell growth, such that after 2.5 h only about a 10 to 15% loss of cell density was anticipated. The samples were injected and kept incubated in 30 ml of boiling absolute ethanol for 5 min and were subsequently kept on ice (0°C) for 15 min. They were then centrifuged (6,000 × g, 20 min, 4°C), and the supernatant was lyophilyzed. The lyophilysate was redissolved in 4 ml of distilled water. Then, 2.8 ml of this solution was transferred to a standard 10-mm NMR tube holding a coaxial 5-mm NMR tube with D₂O and 1 M K¹⁵NO₃ as a chemical shift reference and concentration standard. The ¹⁵N spectra of these cell extracts were run with the following NMR parameters: 70° pulses (15 µs) with a 6-s recycle delay, 20.8-kHz sweep width, 32,768 complex data points, and continuous Waltz-16 composite-pulse broadband proton decoupling (90° ¹H pulse, 200 µs). A total of 4,000 scans per spectrum were averaged. For concentration calibration, 50 mM ¹⁵N-labelled glutamic acid (99% ¹⁵N; Cambridge Isotope Laboratories) was added to a sample obtained 90 min after the addition of the (¹⁵NH₄)₂SO₄, the pH was adjusted to 7.0, and the increase in the ¹⁵N-NMR signal was determined. Thereafter, 25 mM ¹⁵N₂-labelled glutamine (99% ¹⁵N₂; Cambridge Isotope Laboratories) was added to the same sample at a constant pH, and the signal increase of the ¹⁵N and ¹⁵N of the glutamine, as well as of the ¹⁵N of the glutamate, which was resolved from that of the ¹⁵N of the glutamine, was determined. It appeared that the peak area per millimolar concentration of ¹⁵N was equal for the -amino nitrogens of glutamate and glutamine but was 1.5 times larger for the -amino nitrogen of glutamine due to different NOEs. Spectra were routinely processed with 1-Hz line broadening, and peak areas were determined by interactive integration by using the spectrometer software. In cases where it was needed, Gaussian resolution enhancement was used to resolve overlapping peaks. Resonance assignments were made by comparison with literature data (25). After ¹⁵N-NMR analysis, the cell extracts were filtered and the amino acid concentrations were determined by reversed-phase high-pressure liquid chromatography (HPLC) (LC 1090 HPLC; Hewlett Packard, Waldbronn, Germany) after automatic ortho-phtaldialdehyde precolumn derivation. For pool size determinations, the cytoplasmatic volume in the culture was measured by the ³H₂O-¹⁴C-taurine method (38).

(iii) Determination of ammonium ¹⁵N enrichment. Next, 600 µl of the redissolved lyophilysate (see above) was transferred to a standard 5-mm NMR tube, and the pH was adjusted to ca. 1. A single-scan proton NMR spectrum was obtained by using a 90° pulse, an 8-kHz sweep width, and 8,000 complex data points to determine the percent ¹⁵N enrichment of ammonium as described elsewhere (36).

(iv) Calibration of ¹⁵N enrichments in glutamate and glutamine. In order to determine the percent ¹⁵N enrichments in cytosolic glutamine and glutamate at key time points, the amino acids of several cell extracts were fractionated by cation-exchange chromatography on an Ultrapac 11-µm resin column (Pharmacia Biotech GmbH; Freiburg, Germany) (43) with triethylamine buffer (0.2 M, pH 3.2 to 10.5) for elution. Freeze-dried powders of the amino acids were dissolved in 700 µl of D₂O and passed through a 0.2-µm DynaGard filter (Microgon Inc., Laguna Hills, Calif.) to remove insoluble impurities, and the pH was carefully adjusted to 7.0. ¹⁵N enrichments in glutamine and glutamate were determined by ¹H-NMR by using a heteronuclear spin-echo difference spectroscopy protocol adapted from earlier studies (45, 53). The basic pulse sequence schematically reads: ¹H: 90°_X  T_H  (90°_Y  A  90°_Y)  T_H  Acquire
¹⁵N:  T  (90°_X  B  90°_X)  T_N

X

Estimation of the principal ammonium assimilating fluxes. The time-dependent incorporation of ¹⁵N in cytosolic glutamine and glutamate observed with in vivo NMR was described by using a model of differential equations. In this model of the principal ¹⁵N ammonium assimilation pathways in C. glutamicum, the following reactions were included: (i) reversible ammonium diffusion over the cell membrane, characterized by a rate constant K_in for the resulting import of ammonium into the cell and a rate constant K_ex for export from the cell; a constant net uptake of ammonium F₀; the GDH reaction, formulated as a bidirectional flux F_1X superimposed on a unidirectional net flux F₁; the GOGAT reaction, formulated as a bidirectional flux F_2X superimposed on a unidirectional flux F₂; the GS reaction, formulated as an irreversible flux F₃; a net flux of glutamate consumption F₄, accounting for transamination of glutamate in a variety of biosyntheses (e.g., of amino acids as well as for incorporation of glutamate in cell protein and for the build-up of its cytoplasmic pool); a net consumption F₅ of glutamine, accounting for the build-up of its intracellular pool and for incorporation in the cell protein; and a net flux of amidotransferase of glutamine to glutamate F₆, accounting for glutamine functioning as a nitrogen donor in the biosynthesis of, for example, nucleotides, cell wall components, and aromatic amino acids (12). Incorporation of N into the biomass was considered irreversible. The model is schematically shown in Fig. 1.

View larger version (25K): [in this window] [in a new window]   FIG. 1.   Model of the nitrogen fluxes involved in ammonium assimilation in C. glutamicum. This model was fitted to the experimentally determined 15N-NMR data in order to estimate the fluxes in vivo. Each rectangle represents a specific nitrogen atom pool.

	RESULTS

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Activities of ammonium assimilatory enzymes in C. glutamicum during N and C limitation. In continuous fermentations of C. glutamicum the ammonium availability was varied by using the independent NH₄⁺ dosing system according to the following protocol: after an initial growth phase of 40 h, ammonium-limited continuous cultivation was run for 40 h, whereafter the ammonium limitation was relieved and gradually (20 h) changed to C limitation, after which carbon-limited continuous cultivation was continued for another 40 h. The glucose concentration during N limitation was 15 to 20 g/liter, and the NH₄⁺ concentration during carbon limitation was 30 to 50 mM. The specific activities of GS, GDH, and GOGAT were determined in cell extracts taken at several time points during the fermentation. The results are given in Table 2.

Growth to high cell density and mixing time of labeled ammonium. For the present study, we further optimized the combined in vivo NMR-fermentation system until it was suited for the aerobic continuous cultivation of C. glutamicum at 50 g (dry weight)/liter, where in vivo ¹⁵N-NMR spectroscopy could be performed with a time resolution of down to 8 min. The in vivo ¹⁵N experiments were performed on carbon-limited high-cell-density continuous cultures. The wild type reached a final density of 50 g (dry weight)/liter already 15 h after inoculation, whereas the GDH mutant, cultivated with identical fermentation parameters, needed considerably more time, 35 h, to reach the same density (data not shown). This indicates a significant growth advantage of the wild type over the GDH mutant strain. The in vivo experiment was started by the addition of 20 g of labeled (¹⁵NH₄)₂SO₄ after another 50 or 30 h of steady-state continuous cultivation for the wild type and the GDH mutant, respectively. Ideally, mixing of the labeled and unlabeled ammonium should be much faster than the time resolution of the in vivo experiment to avoid an observable influence on the kinetics of label accumulation in the ammonium assimilatory pools. The ¹H-NMR spectra of acidified cell extracts show separate signals of ¹⁴NH₄⁺ and ¹⁵NH₄⁺ and therefore were used to check the kinetics of the mixing of the added labeled ammonium with the unlabeled NH₄ initially present. An example ¹H-NMR spectrum is shown in Fig. 2. The experiments showed that equilibrium ¹⁵N labeling of ammonium was indeed reached within 2 min after application of the (¹⁵NH₄)₂SO₄ pulse (data not shown), illustrating the short mixing time of the fermenter system due to the high recirculation flow of 800 liters/h. Thus, the total ammonium concentration rose within 2 min from 40 mM to ca. 150 mM in the experiments. The final percents ammonium ¹⁵N labeling were 78% in the wild-type experiment and 80% in the experiment with the GDH mutant.

View larger version (13K): [in this window] [in a new window]   FIG. 2.   Example 1H-NMR spectrum of an acidified cell extract obtained 120 min after the addition of 15NH4+ to the C. glutamicum ATCC 13032 culture. The doublet results from 15NH4+ due to scalar coupling of the protons with 15N (spin 1/2); the triplet results from 14NH4+ due to scalar coupling of the protons with 14N (spin 1). These spectra were used to follow the kinetics of the mixing of the added labeled ammonium with the unlabeled ammonium initially present.

In vivo monitoring of ¹⁵NH₄ assimilation. To identify the primary products and follow the kinetics of ammonium assimilation in the wild type and the GDH mutant, in vivo ¹⁵N-NMR spectroscopy experiments were performed on carbon-limited high-cell-density continuous cultivations, one for each strain. The series of in vivo ¹⁵N-NMR spectra obtained with the wild-type strain are shown in Fig. 3. By employing a measurement time of 15 min per spectrum, the signals from the amino nitrogen of glutamate at 335 ppm and from ammonium at 355 ppm could readily be observed already in the first spectrum, identifying glutamate as a prime target of ammonium nitrogen assimilation. However, unexpectedly, the first spectrum also contained a signal from the amido nitrogen of glutamine at 264 ppm, indicating that this compound must play a key role in ammonium assimilation in C. glutamicum ATCC 13032. After ca. 1 h of incubation with the labeled substrate, signals from the amino nitrogen of proline (320 ppm) and from the -amino nitrogen of lysine (343 ppm) appeared. Proline directly derives from glutamate, whereas the lysine nitrogen is labeled via transamination reactions. The late appearance of these compounds in the spectra shows that they do not play a role in primary ammonium assimilation. The same holds true for alanine, of which even in the last spectra no peaks were observed, thus ruling out any role for this amino acid. Whereas alanine was shown to play a major role in nitrogen metabolism in Agaricus bisporus (2), apparently it does not do so in C. glutamicum under the conditions studied. The strong label in the -amino nitrogen of glutamate was in part expected from the high levels of this metabolite known to be present in C. glutamicum (24), but it also points at a significant activity of GDH in vivo since this large pool was labeled to half-maximum already after ca. 20 min. The observed incorporation of ¹⁵N label into glutamine N was surprisingly fast, i.e., achieving half-maximum after ca. 15 min. The relatively strong intensity of this signal suggests that C. glutamicum wild type has, in addition to the large glutamate pool, a very significant glutamine pool as well. Given the large amount of ¹⁵NH₄⁺ added at time zero, the ammonium NMR signal is only weak. Furthermore, it demonstrated a curious behavior: it continuously diminished and changed its sign after ca. 1 h. While this may suggest some uptake phenomenon, it rather reflects a steadily decreasing NOE factor for ammonium during the experiment. Since this factor for ¹⁵N upon proton decoupling is negative, the point of zero-crossing corresponds to an average NOE factor of 1 for ammonium (3).

View larger version (33K): [in this window] [in a new window]   FIG. 3.   Series of in vivo 15N-NMR spectra obtained with C. glutamicum ATCC 13032 before (0 min) and in subsequent 15-min intervals after the addition of [15N]ammonium.

View larger version (36K): [in this window] [in a new window]   FIG. 4.   Series of in vivo 15N-NMR spectra obtained with the GDH mutant of C. glutamicum before (0 min) and in subsequent 7.5-min intervals after the addition of [15N]ammonium.

Compounds observed in extract spectra and deconvolution of overlapping signals. ¹⁵N-NMR spectra of ethanolic cell extracts, due to their superior resolution and signal-to-noise ratio, allowed identification of signals that were not detectable in the in vivo spectra. Figure 5 presents an illustrative cell extract spectrum of a sample taken from the GDH mutant fermentation. Confirming the in vivo measurements (Fig. 4), the most prominent peaks stem from the amido nitrogen of glutamine at 264.3 ppm, from the amino nitrogens of both glutamate and glutamine at 335.5 ppm, and from ammonium at 355.5 ppm. Smaller signals, which were not yet visible in the in vivo spectrum taken at the corresponding time point due to the limited signal-to-noise ratio, are from the amino nitrogen of proline (320.7 ppm), the -amino nitrogen of lysine (343.5 ppm), and the -amino nitrogens of alanine (333.1 ppm) and valine (339.6 ppm). The glutamine signal at 264.3 ppm partially overlaps with two unidentified signals at 264.1 and 264.0 ppm (one of them probably asparagine), and the glutamate-glutamine peak at 335.4 ppm overlaps with two small unidentified signals at 334.9 and 335.9 ppm. Another small signal (tentatively assigned to aspartate [18]) appears at 336.8 ppm. These latter three peaks were not resolved from the glutamate-glutamine peak at 335 ppm in the in vivo spectra. Therefore, their relative areas were determined from the extract spectra and subsequently used to correct the glutamate and glutamine peak areas determined from the in vivo spectra. The linearly increasing relative total overlapping signal for the GDH mutant after 2 h amounted up to ca. 20% at 355 ppm and 7% at 264 ppm, whereas for the wild type it reached 25% at 355 ppm and 15% at 264 ppm.

View larger version (12K): [in this window] [in a new window]   FIG. 5.   Illustrative 15N-NMR spectrum of a cell extract taken from the GDH mutant of C. glutamicum 120 min after the addition of the [15N]ammonium. Assignments are based on the literature (25).

Relative ¹⁵N enrichments and pool sizes indicate a key role of glutamine. The in vivo ¹⁵N data represent only relative peak areas. They must be scaled to absolute intracellular concentrations to enable the further analysis necessary to determine the nitrogen fluxes. This scaling cannot be done in a straightforward manner because the in vivo intensities are strongly mediated by the NOEs that are impossible to predict theoretically or to mimic experimentally by using standards. Therefore, glutamate and glutamine were purified from a number of ethanolic cell extracts taken at representative time points, and the ¹⁵N enrichment in the various positions were determined. For this purpose, heteronuclear ¹H spin-echo difference spectroscopy measurements were performed as described in the Materials and Methods section. The results are given in Table 3.

Fluxes in the ammonium assimilatory pathways. The in vivo ¹⁵N-NMR data, calibrated with absolute pool sizes and ¹⁵N enrichments determined from samples taken at key time points as described above, could now directly be converted to amounts of nitrogen accumulated per gram of biomass and per minute. The differential equation flux model was then fitted to the experimental data of the wild type and the GDH mutant as described above to estimate the model parameters. The results of these fits, which are estimated values for the nitrogen fluxes, are given in Table 4.

View larger version (9K): [in this window] [in a new window]   FIG. 6.   Experimental (squares, N of glutamate and glutamine; circles, N of glutamine) and predicted NMR peak areas for the wild-type ATCC 13032 (A) and the GDH mutant (B) strains corresponding to the model fit results given in Table 4. Peak areas were corrected for overlap by using the 15N-NMR results obtained with ethanolic extracts. The downward trend in the peak area data reflects the dilution of biomass in the fermenter due to sample taking procedures.

	DISCUSSION

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In previous in vivo ¹⁵N-NMR studies on concentrated cell suspensions taken from growing B. lactofermentum cultures, the prominent signals observed were from ¹⁵N-glutamate, ¹⁵N-glutamine, ¹⁵N-alanine, and ¹⁵N-lysine (13). These signals were also observed in the present in vivo ¹⁵N-NMR study on C. glutamicum cultivated at a high cell density under well-defined steady-state conditions in the NMR bioreactor system. However, the strong signal of N-acetylglutamine observed in packed cell spectra of B. lactofermentum (13) was not present in the spectra shown in the present study. Since in that study considerable time was needed to concentrate the cells and to measure the spectra (130- to 200-min data accumulation time per spectrum), it must be concluded that the cells were not in a defined state during those experiments. In experiments mimicking those of Haran et al. (13), signals of N-acetylglutamine were also observed with C. glutamicum. Therefore, the occurrence of this compound in glutamate-producing bacteria might be related to the presence of anaerobiosis or of cell lysis.

Comparison of the ¹⁵N-NMR spectrum of an ethanolic cell extract (Fig. 5) with the corresponding in vivo ¹⁵N-NMR spectra (Fig. 4) shows that the ratio of the glutamine N to the glutamate-plus-glutamine N signal intensity differs strongly in the two cases. This can be explained by a difference in NOE factors in the in vivo versus the extract NMR measurement. The NOE results upon proton broadband decoupling as used in this study from ¹⁵N relaxation by dipolar interaction with protons. Since NOE factors for ¹⁵N are negative and may range from zero to a theoretical maximum of 4.93, the NOE may even cause nulling of signals under specific conditions (3), as was indeed observed in this study (Fig. 3). The NOE factor, when added to the original signal intensity (+1.0), results in a maximum NOE enhancement of 3.93. When the NOE factor is around 1.0, a strong sensitivity of the observed signal towards this factor results since, for example, a minor increase in NOE factor from 1.1 to 1.2 would produce a doubling of the observed ¹⁵N-NMR signal from 0.1 to 0.2 (relative intensity). Paramagnetic ions exert a significant influence on the ¹⁵N NOE by their contributing to the relaxation of the ¹⁵N nucleus. This can lead to a variable behavior of ¹⁵N signal intensities depending on environmental conditions such as medium mineral composition and cell density (1, 15) as observed in this study (cf. Fig. 3, 4, and 5). Most likely, the fact that paramagnetic ions bind much more easily to the -amino (and the carboxylate) than to an amide group causes a reduction of the NOE factor for the former in vivo.

The specific activities of the NH₄⁺-assimilating enzymes in the wild type under conditions of NH₄⁺ abundancy as determined in this study are comparable to previously reported values (5, 41, 46, 49). The specific activities of GDH and GOGAT determined under nitrogen limitation conditions in this study were likewise comparable to values measured at low ammonium concentrations (11, 50). However, the specific GS activity found under N-limiting conditions (ca. 11 U/mg of protein) was 5- to 10-fold higher than that reported in comparable studies (47, 49). This could be a consequence of the fact that in none of those studies did C. glutamicum grow strictly in ammonium-limited conditions with carbon abundance, as was the case in the present work.

The observed regulatory behavior of GS and GOGAT in C. glutamicum ATCC 13032 in this study seems to be common to many microorganisms possessing both GDH and GS-GOGAT (33) and is similar to that observed by Kim et al. (21) and Sung et al. (47) for C. glutamicum ATCC 13058 and Brevibacterium flavum. The results of the present study therefore are consistent with GS being regulated directly by the nitrogen availability in the medium. The regulation mechanism possibly proceeds via adenylation of GS at high NH₄⁺ concentrations (35), a finding comparable to the known mechanism in Escherichia coli (37).

The observed weak dependency of GDH on the ammonium availability in the medium corroborates the finding of Sung et al. (47) for GDH in B. flavum but differs from that found with C. glutamicum ATCC 13058, where the GDH activity significantly decreased with increasing NH₄⁺ concentrations (21).

The in vivo NMR experiments presented in this study revealed that the glutamine pool was the second largest cytoplasmic amino acid pool in wild-type C. glutamicum ATCC 13032. Since it was observed that glutamine was easily hydrolyzed in an acidic environment (data not shown), the most likely reason that the important presence of glutamine in this bacterium has remained undetected thus far is because most studies of intracellular metabolites used perchloric acid extraction (10), thereby hydrolyzing the glutamine to glutamate. Differences in employed ammonium concentrations are unlikely to have caused elevated glutamine levels, since the steady-state ammonium concentration in our study (ca. 40 mM) was not very much different from that employed in most previous studies (typically 20 to 40 mM). Since, moreover, these concentrations are well in excess of the K_m values of GDH and GOGAT for NH₄⁺, it is also unlikely that even the 100 mM ammonium pulse applied in the present study has influenced the glutamate and glutamine pool sizes.

The flux analysis revealed the surprising fact that during chemostat growth under carbon limitation and NH₄⁺ abundance, nearly 28% of the NH₄⁺ was assimilated via the GS reaction in glutamine, while 72% was assimilated by the GDH reaction via glutamate. Thus, this study uncovered a previously unrecognized, important role of glutamine in NH₄⁺ metabolism of C. glutamicum. The value of 28% is approximately twice as high as the glutamine requirement for biomass synthesis in E. coli, which was reported to be 13% of the total assimilated nitrogen (37). This difference can be explained by the different cell wall compositions of gram-positive C. glutamicum and gram-negative E. coli. Glutamine functions as a nitrogen donor in the synthesis of carbamoyl phosphate, histidine, purines, and glucosamine-6-phosphate, a precursor of peptidoglycan (12). The fact that the amount of peptidoglycan in gram-positive bacteria is three- to sevenfold higher than that of gram-negative bacteria (39) may thus well be responsible for the strong use of glutamine for biomass synthesis of C. glutamicum observed in the present study. In fact, by using published data on C. glutamicum biomass composition (30) and taking into account the higher peptidoglycan content that can be calculated from the diaminopimelate content (146 µmol/g [dry weight]) of C. glutamicum, the amount of glutamine needed for the synthesis of a 1-g biomass can be calculated as ca. 2,000 µmol/g (dry weight). Glutamine amidotransferases make up ca. 1,800 µmol/g (dry weight) of this amount. The total requirement of nitrogen can likewise be calculated from the data of Marx et al. (30) modified for the higher peptidoglycan content; it amounts to ca. 9,500 µmol/g (dry weight). Thus, according to this calculation, 21% of the total nitrogen is assimilated via glutamine by direct incorporation or the above-mentioned glutamine amido-transferring enzymes (represented by F₅ and F₆, respectively, in our flux model [Fig. 1]). This compares rather well with the value of 28% found in the present in vivo study (Table 4).

The specific GOGAT activity in the GDH mutant C. glutamicum was relatively high and, in marked contrast to the wild type, showed only a weak dependency on the availability of NH₄⁺ (Table 2). This was also observed by Börmann-El Kholy (5). A comparably altered regulatory behavior of GOGAT has also been described for a GDH defect mutant of B. flavum (47). Since the GOGAT activity in the GDH mutant of C. glutamicum was essentially independent of ammonium availability in the medium, its regulation mechanism must be different from that of GS. The fact that the glutamate pool in the mutant was found to be much lower than in the wild type in the present study suggests that a reduced glutamate concentration may provide the signal for induction of GOGAT expression in C. glutamicum. The observation that glutamate or glutamate-generating nitrogen substrates (arginine, proline, or histidine) repress GOGAT expression in Salmonella typhimurium and Klebsiella aerogenes during N-limited growth (6, 7) is in agreement with this hypothesis. The possibility that the observed elevated GOGAT activity in the GDH mutant at high extracellular NH₄⁺ concentrations might be a consequence of a severe limitation in ammonium transmembrane transport in this strain can be definitively ruled out. Indeed, the flux analysis performed in this study indicated that a very fast equilibration of ammonium across the C. glutamicum membrane took place, i.e., the initial concentration gradient of 100 mM was compensated within 2 min. Literature data on this subject are scarce. While the ammonium uptake system in C. glutamicum has been characterized (23, 24), this system is not expressed at high ammonium concentrations as pertinent in the present work. Therefore, net ammonium uptake must be a consequence of ammonia diffusion across the cell membrane. A review (22) mentions ammonia permeability coefficient values for Nitella clavata (9 · 10⁶ m/s) and Klebsiella pneumoniae (5 · 10⁷ m/s). Using the average of the two values (4.75 · 10⁶ m/s) for the permeability coefficient, assuming a volume-to-area ratio of 2 · 10⁷ m for C. glutamicum (modelled as an ellipsoid with axes of 2 and 1 µm), taking into account that at neutral pH the equilibrium ratio of the NH₄⁺ and NH₃ concentrations is 180 and applying Fick's First Law of Diffusion, we obtain d[NH₄⁺]_in/dt = (60 · 4.75 · 10⁶) · ([NH₄⁺]_ex  [NH₄⁺]_in)/(2 · 10⁷ · 180), from which a time constant for the equilibration of ammonium across the cell membrane of 0.13 min can be calculated. This agrees well with the value estimated by the flux modelling, given the uncertainty of the applied value for the permeability coefficient.

The results of the in vivo ¹⁵N-NMR nitrogen flux analysis of the GDH mutant of C. glutamicum were completely in accordance with the pattern expected from the enzyme data (Table 2), i.e., the absence of GDH flux, the high activity of GS, and the glutamate synthesis by operation of GOGAT. This demonstrates the reliability of the present approach. Again surprising was the even higher flux over glutamine amidotransferase fluxes (F₆), which amounted to almost 43% of total assimilated nitrogen (Table 4). The confidence region of this glutamine amido-transferring flux estimation, however, is rather large because the estimation of F₆ was highly correlated with that of the GOGAT flux F₂ which showed a moderately high standard deviation (Table 4). Thus, a fraction of 25 to 30% of the total assimilated nitrogen, which was also found for the wild type and which is not very much different from the calculated value of 21%, does not contradict the glutamine amidotransferase flux estimate for the GDH mutant. Our observation that the GOGAT reaction was partly reversible (Table 4) is at variance with the irreversibility demonstrated for the enzyme (referred to as glutamate synthase) from Azospirillum brasilense (52). However, in that study it was mentioned that the A. brasilense enzyme differed from the glutamate synthases from K. aerogenes and E. coli, which were weakly reversible, and the C. glutamicum enzyme may likewise be different in this respect. Moreover, the precision of our estimate does not allow us to fully exclude an irreversibility of the C. glutamicum GOGAT in vivo.

We observed that the GDH mutant grew much slower to high cell density than the wild type. This observation, together with the flux analysis results, identifies GOGAT as the growth-limiting bottleneck in the GDH mutant. This phenotype remained concealed in strain characterization studies of this mutant by Börmann-El Kholy (5), possibly because this author used a combination of ammonium and urea as nitrogen source instead of only NH₄⁺ as in the present study. This difference in nitrogen source composition may have influenced the regulation state of the NH₄⁺-assimilating enzymes in the two C. glutamicum strains.